bioRxiv preprint doi: https://doi.org/10.1101/2020.10.21.345918; this version posted October 21, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-NC-ND 4.0 International license.

The molecular coupling between substrate recognition and ATP turnover in a

AAA+ hexameric loader

Neha Puri1,2, Amy J. Fernandez1, Valerie L. O’Shea Murray1,3, Sarah McMillan4, James

L. Keck4, James M. Berger1,*

1Department of Biophysics and Biophysical Chemistry, Johns Hopkins School of

Medicine, Baltimore, MD 21205

2Bristol Myers Squibb, 38 Jackson Road, Devens, MA 01434

3Saul Ewing Arnstein & Lehr, LLP, Centre Square West, 1500 Market Street, 38th Floor,

Philadelphia, PA 19102

4Department of Biomolecular Chemistry, University of Wisconsin School of Medicine

and Public Health, Madison, WI, 53706

*Corresponding author

Email: [email protected]

Keywords: DNA replication, AAA+ ATPase, Helicase, Meier-Gorlin Syndrome

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ABSTRACT

In many and in eukaryotes, replication fork establishment requires the

controlled loading of hexameric, ring-shaped around DNA by AAA+ .

How loading factors use ATP to control helicase deposition is poorly understood. Here,

we dissect how specific ATPase elements of E. coli DnaC, an archetypal loader for the

bacterial DnaB helicase, play distinct roles in helicase loading and the activation of DNA

unwinding. We identify a new element, the arginine-coupler, which regulates the switch-

like behavior of DnaC to prevent futile ATPase cycling and maintains loader

responsiveness to replication restart systems. Our data help explain how the ATPase

cycle of a AAA+-family helicase loader is channeled into productive action on its target;

comparative studies indicate elements analogous to the Arg-coupler are present in

related, switch-like AAA+ that control replicative helicase loading in eukaryotes,

as well as clamp loading and certain classes of DNA transposases.

INTRODUCTION

The proliferation of all cells relies on the accurate and timely copying of DNA by large

macromolecular assemblies termed . Cellular replisomes are complex, multi-

functional assemblies that co-localize a number of essential biochemical functions to

increase replication efficiency and speed (O’Donnell et al.). One such function is the

unwinding of parental to expose template DNA for leading and lagging

strand synthesis, a process carried out by specialized, ring-shaped known as

helicases. In Gram-negative bacteria, an known as DnaB (DnaC in Gram-

positive organisms) serves as the principal helicase for driving replication fork

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progression (reviewed in Chodavarapu and Kaguni, 2016). DnaB proteins form

hexameric rings and are generally incapable of spontaneously associating with

chromosomal replication start sites (origins). As a consequence, DnaB helicases are

specifically recruited and deposited onto single-stranded origin DNA by dedicated

initiation and loading factors.

Replicative helicase loading in bacteria depends on at least one of two known types of

proteins. One widespread class, comprising factors related to a known as DciA,

is ATP-independent (Brézellec et al., 2016; Mann et al., 2017). The second class is

composed of DnaC/DnaI-family proteins, which are ATP-dependent and present in

model organisms such as E. coli and B. subtilis (Brézellec et al., 2016). Both DnaC and

DnaI consist of an N-terminal helicase-binding domain (HBD) fused to a C-terminal

AAA+ (ATPases Associated with various cellular Activities) nucleotide-binding fold (Fig.

1A) (Ludlam et al., 2001; Neuwald et al., 1999b). The AAA+ region of DnaC contains

numerous signature sequence motifs (Walker A, Walker B, Sensor I & II, Arginine-

finger), several of which have been reported to be critical for the function of the loader in

E. coli (Makowska-Grzyska and Kaguni, 2010; Mott et al., 2008). A few of these

elements have been examined and found to be important for controlling aspects of

DnaC activity in vitro as well (Davey et al., 2002; Makowska-Grzyska and Kaguni, 2010;

Mott et al., 2008) however, a systematic analysis of their individual mechanistic impact

on the efficiency of DNA loading and unwinding by DnaB has been lacking.

3 bioRxiv preprint doi: https://doi.org/10.1101/2020.10.21.345918; this version posted October 21, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-NC-ND 4.0 International license.

DnaC-type loaders help place DnaB hexamers on single-stranded DNA by opening the

helicase ring and allowing a nucleic acid strand to enter laterally into the pore of the

complex (Arias-Palomo et al., 2019). In the context of DnaB, six DnaC protomers form a

cracked hexameric oligomer, generating composite ATPase active sites at five of the six

subunit interfaces (Fig. 1B,C) (Arias-Palomo et al., 2013, 2019). The AAA+ fold of

DnaC binds ssDNA directly, at a site distal from its ATP-binding locus (Arias-Palomo et

al., 2019; Ioannou et al., 2006; Mott et al., 2008). Prior to binding DNA, six DnaC

protomers latch onto a DnaB hexamer through their HBDs to promote helicase ring

opening. In this configuration, EM studies have shown that DnaB resides in an ADP-like

state, and DnaC in an ATP-bound form (Arias-Palomo et al., 2019). DNA binding

induces a large conformational rearrangement in the DnaB hexamer and compresses

the DnaC ATPase ring, switching the ATPase status of the helicase and the loader into

ATP and ADP-forms, respectively. Curiously, although DnaB helps promote the local

co-association of neighboring DnaC AAA+ domains, the ATPase activity of the loader

does not manifest until ssDNA is bound (Arias-Palomo et al., 2013; Davey et al., 2002;

Kobori and Kornberg, 1982) How DnaB promotes DnaC self-assembly yet does not

directly trigger ATPase turnover by the loader is not understood. How DNA binding is

relayed to the DnaC ATPase site and whether DnaB ATPase function is important for

loading (as structural work suggests) are similarly unknown. Addressing these

questions is important for understanding how replicative helicase loaders exploit

nucleotide-coupling mechanisms to generate productive action on client substrates, a

question relevant to AAA+ ATPases in general.

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Here, we show through a structure-guided biochemical approach that defects in

different DnaC ATPase elements have distinct impacts on the ability of the loader to

deposit DnaB onto DNA and to activate helicase unwinding of forked substrates. DnaC

is shown to be capable of promoting helicase unloading in an ATP-dependent manner;

both loading and unloading are additionally found to be influenced by the ATPase

activity of the helicase as well. We identify a set of residues in DnaC that couple loader

ATPase activity to the binding of DnaB and ssDNA and that unexpectedly modulate the

responsiveness of DnaC to a replication restart factor, and we show that analogs of one

of these elements, an active-site arginine, are present in related AAA+ superfamily

members that operate with switch-like properties. Our data collectively explain how the

ATPase status of DnaC is coupled to recognition of its DnaB and ssDNA substrates to

ensure tight coupling of helicase loading and DNA unwinding with nucleotide turnover.

RESULTS:

Conserved AAA+ motifs support DnaC ATPase activity

We first set out to address the extent to which five canonical AAA+ motifs support the

ATP-hydrolysis activity of E. coli DnaC. Using site-directed mutagenesis, we generated a

panel of individual DnaC variants lacking one of the five key residues found in AAA+

proteins (Walker A – K112R; Walker B – E170Q; Sensor I – N203A; Arg-finger – R220D;

Sensor II – R237D). We expressed and purified each of these mutants and, using a

coupled assay, tested them for ATP hydrolysis activity in the presence of a hydrolysis

defective DnaB mutant (DnaBKAEA) and ssDNA. The ablation of any of the five residues

was found to result in complete loss of ATPase function (Fig 2A). These results highlight

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the importance of the AAA+ signature residues to the nucleotide turnover capabilities of

E. coli DnaC, and further establish that the hydrolysis activity present in the assay derives

from the loader and not the helicase or a contaminating ATPase.

DnaC ATPase function is coupled to helicase loading efficiency

Since the ATPase activity of DnaC requires the presence of DnaB (Arias-Palomo et al.,

2019; Davey et al., 2002) we next asked how the ATPase cycle of the loader contributes

to productive interactions with its client helicase. The role of each DnaC AAA+ ATPase

residue was investigated in the loading of DnaB onto a closed-circular,

M13mp18 ssDNA substrate using a gel-filtration chromatography assay that resolves

ssDNA-DnaB-DnaC complexes, free DnaB, and DnaC (Davey et al., 2002). As controls,

efficient DnaB loading was observed in the presence of wild type DnaC (Fig. 2B),

whereas little ssDNA association was detected for DnaB in absence of DnaC (Fig. S1A).

By comparison, the loading efficiency of the DnaC HBD was approximately half that of

WT DnaC (Figs. 2C, S1B). Investigation of the remaining active site mutants show that

they had a variable effect on helicase loading efficiency, with the activity of the Sensor I

mutant proving the most compromised (slightly moreso than the HBD alone) (Figs. 2C,

S1C). The loading activities of the Walker-B and Arg-finger mutants were roughly

commensurate with the HBD, whereas the Sensor II and Walker-A mutants were

effectively wild-type (Figs. 2C, S1D-G). Collectively, these results demonstrate that the

AAA+ domain and the ATP-hydrolysis cycle of DnaC increase the efficiency of helicase

loading. They also show that, despite being required for ATPase function overall, different

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AAA+ active site residues contribute to the efficiency of DnaB deposition to varying

degrees.

Helicase loading is aided by DnaB’s ATPase activity and results in the topological

entrapment of ssDNA

Prior structural work has indicated that DnaC-associated DnaB switches from an ATP to

an ADP state upon binding DNA, and that the engagement of nucleic acid triggers DnaB

to adopt a cracked-ring state that is topologically linked to its DNA substrate through

domain swapping interactions (Arias-Palomo et al., 2019). To better define the extent to

which the ATPase activity of DnaB is important for loading, we assessed the ability of

wild-type DnaC to load either native DnaB or a double Walker-A/B ATPase-defective

mutant (DnaBKAEA) of the helicase using the M13 association assay. The resultant data

show that the ATPase deficient DnaB is loaded by DnaC, but at a reduced level (~50%)

compared to the wild type helicase (Figs. 3A, S2A). We next increased the concentration

of salt in the sizing column running buffer used to separate DNA-loaded and -free DnaB.

A majority of the loaded DnaB remained associated with ssDNA when NaCl was

increased from 100 to 250 mM NaCl (Figs. 3B, S2B). These results demonstrate that the

ability of DnaB to hydrolyze nucleotide is not critical for loading, but does aid the efficiency

of the deposition reaction, and that once loaded, DnaB persists in a salt-stable complex

with ssDNA. This latter finding indicates that the topologically closed-ring conformation

observed structurally for the DnaBC complex on DNA persists in solution.

DnaC can promote helicase unloading

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Wild-type DnaC is known to remain associated with DnaB following loading (Davey et al.,

2002; Makowska-Grzyska and Kaguni, 2010). However, our steady-state ATPase assays

show that DnaC also continuously hydrolyzes nucleotide in the presence of DNA,

indicating that the loader can cycle back to an ATP-bound state. This behavior suggested

that when associated with DnaB and ssDNA, ADP-bound DnaC might be capable of

switching back to an ATP-state and reopening the helicase ring to promote helicase

unloading. To test this idea, we repeated our M13 ssDNA-association reactions with wild-

type DnaC, but now using sizing columns pre-equilibrated with ATP or ADP. The

presence of ADP in the column buffer still resulted in efficient DnaB loading. By contrast,

the inclusion of ATP led to a near complete loss of DnaB that remained associated with

M13 DNA (Figs. 3C, S2C-D), demonstrating that DnaC can unload DnaB in an ATP-

dependent manner. To determine whether the ATPase activity of DnaB might also play a

role in unloading, we assessed the effect of ATP in the column running buffer using the

DnaBKAEA mutant. Interestingly, substantially more ATPase-deficient DnaB remained

loaded on the M13 ssDNA in presence of ATP compared to wild-type DnaB (Figs. S2E-

F). This finding shows that the ATPase activity of DnaB is important for its unloading from

DNA, as well as for loading.

Helicase activation is modulated by DnaC ATPase activity

In addition to controlling DnaB loading, DnaC has been shown to stimulate the unwinding

of forked DNA substrates by the helicase (Arias-Palomo et al., 2013). Unlike closed-

circular M13 ssDNA, DnaB can thread onto free ssDNA ends to promote strand

separation of downstream duplexes without assistance (Jezewska et al., 1998); however,

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unwinding is inefficient unless DnaB is present in large excess or DnaC is included (Arias-

Palomo et al., 2013; Galletto et al., 2004a; Kaplan, 2000; Kaplan and Steitz, 1999). To

define the role of the DnaC ATPase in promoting DnaB helicase activity, we conducted

DNA unwinding assays with our panel of mutants using a forked substrate bearing a Cy3

fluorophore quenched with BHQ2 at the duplex end. When present at only a 2-fold molar

excess over the forked substrate, DnaB was observed to unwind ~10% of the substrate

(Fig. 4A). To determine whether DnaB was inactive in this assay because a fraction of

the helicase associates with the 3’ tail of the fork and impedes access to the 5’ tail, we

blocked the 3’ end of the fork with a biotin-neutravidin complex and repeated the assay.

This block resulted in a higher proportion of observed fork unwinding, but the total amount

of unwound product (~45%) was still less than seen when DnaC was present (~80%)

(Fig. 4A). Interestingly, the level of stimulation afforded by DnaC is comparable whether

a 3’-end block is present or not (compare “B + C” curves in Figs. 4A vs. 4B). These results

indicate that 3’ DNA ends can sequester or inactivate a proportion of DnaB, but that DnaC

can overcome this inhibition.

The ability of different DnaC ATPase mutants to activate helicase unwinding was tested

next. As seen previously (Arias-Palomo et al., 2013), the isolated HBD of DnaC readily

stimulates DnaB activity on an unblocked fork, at a level close to that of the full-length

loading factor (Fig. 4B). By comparison, the DnaC ATPase mutants were much less

capable in their response (Figs. 4C, S3). For example, the Walker-A mutant showed ~40-

50% the stimulatory activity of wild-type DnaC, whereas the charge-reversed Arg-finger

mutant failed to stimulate DnaB beyond the basal level of the helicase alone (other

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mutants were intermediate in their response, with ~25-30% the activity of the native

loader). There was no clear trend between the degree to which a particular mutation

disrupted both DnaC ATPase activity and stimulation of DnaB unwinding. However, the

one mutant that would be expected to interfere with ATP binding (Walker-A) evinced the

highest stimulatory effect, while the mutant that would be most expected to impede inter-

subunit communication (Arg-finger) was the most severely compromised. This behavior

suggests that the capacity of adjoining DnaC protomers to interact with one another in a

nucleotide dependent manner can influence the extent to which the loader can stimulate

DNA unwinding by DnaB.

Multiple sensors link ssDNA binding to DnaC ATPase activity

Although AAA+ ATPase activity is often tightly coupled to interactions with client

substrates, the mechanisms behind this linkage are highly varied and frequently unclear.

Recent structural studies of DnaC bound to DnaB and ssDNA noted three conserved

amino acids – Phe146, Ser177, and Tyr179 – which associate with the nucleic acid strand

(Figs. 1, 5A) (Arias-Palomo et al., 2019). Mutation of these residues blocks ssDNA

binding and DNA-stimulated ATPase activity, as well as the loading of DnaB onto M13

DNA (Arias-Palomo et al., 2019). Their alteration also completely inhibits the ability of

DnaC to stimulate DNA unwinding by DnaB (Figs. S4A-B).

In the course of our phylogenetic analyses of DnaC/I-family proteins, we noted another

conserved residue, Lys143, that is positioned one helical turn upstream of Phe146 (Figs.

1, 5A). Inspection of the DnaBC•ssDNA structure shows that this amino acid is proximal

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to the substrate phosphodiester backbone in a subset of DnaC protomers, forming a small

patch with its Phe146, Ser177, and Tyr179 counterparts that grasps the nucleic acid

substrate (Fig. 5B). Substitution of Lys143 with leads to a complete loss of

DNA/DnaB stimulated ATPase activity by DnaC, and strongly impaired DNA binding

(Figs. 5C, S4C). The K143E mutant also proved incapable of loading DnaB onto M13

DNA or activating DNA strand separation by the helicase (Figs. 5D-E, S4D-E). These

findings demonstrate that, as with other conserved amino acids in its local neighborhood,

Lys143 is a part of the mechanism used by DnaC to sense ssDNA.

A second conserved element in DnaC/I-family proteins revealed by sequence homology

is the presence of an invariant arginine, Arg216, one helical turn upstream of the Arg-

finger (Fig. 1, 6A). To more deeply probe the role of Arg216, we mutated the amino acid

to both aspartate (charge reversal) and alanine (charge ablation) and assessed the

function of these constructs biochemically. The R216D mutant proved unable to hydrolyze

ATP, did not stimulate DNA unwinding by DnaB, and could only support DnaB loading at

level comparable to the Sensor I mutant (Figs. 6B-D, S5A-B). By contrast, DnaCR216A

showed a moderate capacity to support helicase loading and stimulated DNA unwinding

by DnaB with wild-type efficiency (Figs. 6C-D, S5A,C). Surprisingly, the R216A construct

actually exhibited slightly higher ATPase activity than native DnaC and additionally

required only DnaB (and not ssDNA) for ATPase function (Fig. 6B). The alanine mutant

also proved capable of counteracting the strong inhibitory block placed on DnaC ATPase

activity in the context of a K143E substitution (Fig. 6E), although it was unable to restore

the activation of DNA unwinding by DnaB in such a context (Fig. S5D). These results

11 bioRxiv preprint doi: https://doi.org/10.1101/2020.10.21.345918; this version posted October 21, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-NC-ND 4.0 International license.

reveal that Arg216 is as a key coupling element in DnaC, repressing nucleotide turnover

in the presence of DnaB until ssDNA also associates with the complex.

A replication restart mutant disrupts DNA/ATPase coupling in DnaC

In addition to supporting helicase loading during replication initiation, DnaC also

participates in replication restart (Schekman et al., 1975; Wickner and Hurwitzt, 1974).

Numerous DnaC variants have been isolated as suppressors for the loss of critical

replication restart proteins in genetic screens (Sandler et al., 1996); one is dnaC810,

which results from the substitution of Glu176 with glycine. We noted that Glu176, though

not particularly well-conserved, sits proximal to the DNA binding residues Ser177 and

Tyr179, in a loop adjoining the Walker-B element that undergoes a conformational switch

between DNA-free and -bound states of DnaC (Fig. 1C).

To better understand the physical consequences of the dnaC810 mutation, we purified a

E176G version of the protein and assessed its impact on DnaC function in vitro. The

variant proved capable of supporting both DnaB loading and the activation of fork

unwinding by the helicase, at levels only moderately decreased from native DnaC (Figs.

7A-B, S5E-F). DnaCE176G was also able of supporting the ssDNA- and DnaB-dependent

ATPase function of the loader (Fig. 7C); interestingly, the activity of the mutant loader

exceeded that of wild-type protein by ~50%. In addition, the substitution allowed DnaC to

hydrolyze ATP when bound to DnaB alone, although at approximately half the rate as

when DNA was present. These data demonstrate that, like the R216A (Arg-coupler)

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substitution, the mutation present in the dnac810 strain has gained the ability to act on

DnaB in a DNA-independent manner.

The similar biochemical behaviors of the DnaC810 mutant to the R216A variant raised

the question as to whether the Arg-coupler is important for DnaC to bypass the need for

replication restart factors in loading DnaB onto SSB-bound DNA. To address this

question, we tested the ability of DnaCR216A to support DNA unwinding by DnaB on an

SSB-coated fork substrate (Figs. 7D, S5G). Wildtype DnaC only weakly supported fork

unwinding but was modestly stimulated by the presence of the restart factor, PriC. By

contrast, DnaC810 supported a high level of fork unwinding even in the absence of PriC.

Interestingly, DnaCR216A displayed an even greater ability to stimulate DnaB-mediated

unwinding of the SSB-bound substrate than DnaC810 when PriC was absent. These data

demonstrate that the Arg-coupler plays a role in preventing DnaC from activating DnaB

unless restart factors are present.

DISCUSSION

Cells rely on ATP-dependent protein/protein and protein/nucleic-acid complexes to

support a diverse array of critical biochemical processes. A group of enzymes known as

AAA+ ATPases are exemplars of such molecular ‘machines,’ participating in events

ranging from DNA replication and gene expression to proteolytic homeostasis and

vesicle trafficking (Neuwald et al., 1999b). How AAA+ enzymes couple ATP turnover to

movement and function is a broadly studied mechanistic question. Although the

appropriate engagement of client substrates is clearly important for ATPase coupling,

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the elements responsible for these interactions are typically far removed (20-30 Å) from

the nucleotide hydrolysis center. Although there must exist structural linkage elements

that relay information over a distance between client occupancy and ATPase status, the

physical identities of these coupling agents are rarely known.

Here, we have used structure-guided mutagenesis and biochemical analyses to dissect

how DnaC, a bacterial AAA+ ATPase that supports the deposition of the replicative

DnaB helicase onto DNA, coordinates its ATPase cycle with helicase loading and

activation. The opening of the DnaB ring and its subsequent engagement with DNA can

occur independently of ATP hydrolysis by DnaC (Fig. 2), a finding in accord with

previous data showing that neither ATP binding nor the DnaC ATPase domains are

required for DnaB loading in vitro (Arias-Palomo et al., 2013; Davey et al., 2002).

However, ATP turnover does improve the efficiency of helicase loading (Fig. 2), an

observation in agreement with early work of Kornberg and colleagues reporting that an

ATP-hydrolysis step is important in the binding of the DnaBC complex to DNA (Wahles

et al.). Interestingly, the ability to hydrolyze nucleotide also has a potentiating effect on

the capacity of DnaC to activate DNA unwinding by DnaB (Fig. 4). Indeed, hydrolysis-

defective DnaC mutants are less capable of activating DnaB than either an ATP-binding

(Walker-A, K112R) mutant or the helicase-binding domain of DnaC alone. This

observation suggests that, following binding of DnaC to DnaB, ATP turnover by the

loader is important for allowing DnaB to shift into a translocation-competent form.

14 bioRxiv preprint doi: https://doi.org/10.1101/2020.10.21.345918; this version posted October 21, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-NC-ND 4.0 International license.

Recent elucidation of DNA-free and DNA-bound structures of the DnaBC complex have

revealed that the nucleotide status of the helicase flips from an ADP state to an ATP

configuration when DNA is present (Arias-Palomo et al., 2019). This observation

suggested to us that the loading of DnaB onto DNA might be impacted by the helicase’s

ATPase function. In testing this idea with a double Walker-A/Walker-B (KAEA) mutant of

DnaB, the ability of the helicase to interact productively with nucleotide indeed impacts

DNA loading (Fig. 3A). Although this coupling is not a strict dependency, a role for

nucleotide turnover by DnaB is reminiscent of the need for ATP by the eukaryotic

Mcm2-7 replicative helicase to supporting its loading onto DNA by the origin recognition

complex (ORC) (Coster et al., 2014). Interestingly, we find that DnaC can also drive

helicase unloading as well as loading (Fig. 3C). In contrast to eukaryotes, where

unloading of the Cdc45/Mcm2-7/GINS helicase is facilitated by ubiquitylation of MCM7

and removal of the CMG helicase by CDC48/p97 (Deng et al., 2019) relatively little is

known about how DnaB helicases are removed from DNA following replication

termination in bacteria. Additional studies will be needed to assess a potential role for

DnaC in this process.

Structural work has shown that the AAA+ fold of DnaC associates with DNA directly

(Arias-Palomo et al., 2019). Three residues – Phe146, Ser177, and Tyr179 – form a

small DNA-binding locus on the part of the DnaC ATPase domain that faces the interior

pore of the dodecameric DnaBC complex; mutation of these residues ablates DnaB

loading by DnaC (Arias-Palomo et al., 2019). In examining this region further, we found

a fourth amino acid, Lys143, that also participates in binding DNA. Lys143 is generally a

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or arginine in DnaC/DnaI homologs, and a prior genetic screen by Kaguni and

colleagues noted that the integrity of this residue is important for E. coli viability (Hupert-

Kocurek et al., 2007). Here, we show that Lys143 is important for supporting the

ATPase and DnaB-loading activities of E. coli DnaC (Fig. 5); the inability of a DnaCK143E

mutant to facilitate DnaB loading onto ssDNA – even though it has an intact HBD and

stably binds DnaB – suggests that the alteration allows the loader to maintain a

persistent, open-ring state of the helicase. In addition, we find that Lys143 and other

amino acids in the DNA binding patch are critical for DnaC to stimulate fork unwinding

by DnaB (Fig. 5E, S4A-B). Biochemical studies show that disruption of this residue and

other amino acids in the region block DNA binding (Fig. S4A) (Arias-Palomo et al.,

2019) thereby short-circuiting the ability of the nucleic acid substrate to stimulate ATP

hydrolysis by DnaC when the loader is bound to DnaB. Overall, the integrity of the DNA-

binding patch on DnaC supports the ability of ssDNA to activate all known DnaC

functions, even though it is situated over 20 Å away from the DnaC ATPase center.

A long-standing mechanistic question has been to understand why DnaB is required for

stimulating ATP hydrolysis by DnaC, yet is incapable of supporting this activity unless

ssDNA is also present (Davey et al., 2002). Unlike a majority of AAA+ ATPases, E. coli

DnaC is a monomer on its own (Galletto et al., 2004b), and forms stable oligomeric

interactions only when templated by a pre-existing DnaB hexamer (Kobori and

Kornberg, 1982). Formation of a higher-order oligomer state is generally a prerequisite

for AAA+ ATPase function, as the active site of one subunit typically depends on the

contribution of an arginine or lysine from a partner subunit to support nucleotide

16 bioRxiv preprint doi: https://doi.org/10.1101/2020.10.21.345918; this version posted October 21, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-NC-ND 4.0 International license.

turnover (Neuwald, 1999a; Neuwald et al., 1999b). Prior work has shown that mutation

of the so-termed “Arg-finger” residue to alanine in E. coli DnaC does not interfere with

the ability of the loader to deliver DnaB to oriC, but does interfere with DnaC-dependent

DNA replication in vitro and DnaC function in vivo (Makowska-Grzyska and Kaguni,

2010). Here we show that this amino acid is critical for DnaC ATPase activity (Fig. 2A),

and that while it is only moderately important for DnaB loading, its presence is

necessary to activate DNA unwinding by the helicase (Fig. 2C, 4C).

Interestingly, DnaC possesses a second arginine (Arg216) just upstream of its Arg-

finger (Arg220). This amino acid is essentially invariant among DnaC/DnaI homologs

(Fig. 6A). Prior studies have shown that the integrity of this residue is unnecessary for

DnaC to bind ATP or associate with DnaB, but is important for supporting replication in

vitro and for cell viability (Makowska-Grzyska and Kaguni, 2010). Here, we show that

Arg216 is actually an essential element for coupling substrate DNA binding to loader

ATPase activity. Mutation of the amino acid to aspartate severely impairs DnaC’s

ATPase activity and its ability to mobilize and activate DnaB on DNA. However,

substitution with alanine not only increases the rate of ATP hydrolysis by DnaC, but also

allows nucleotide turnover to occur even in the absence of DNA (Fig. 6B). The

DnaCR216A mutant also overrides the ATPase defect imparted by a DNA binding mutant

(e.g., K143E), yet does not degrade the ability of DnaC to assist with either DnaB

loading or the stimulation of DnaB-mediated fork unwinding. Inspection of a DNA-free

DnaBC structure reveals that Arg216 is positioned where the nucleophilic water would

be expected to reside prior to ATP hydrolysis; in the presence of ssDNA, this amino

17 bioRxiv preprint doi: https://doi.org/10.1101/2020.10.21.345918; this version posted October 21, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-NC-ND 4.0 International license.

acid shifts away to allow access to the reactive g-PO4 (Fig. 8A-B, Movie S1). Thus,

Arg216 acts not just as a substrate/ATPase coupler, but also as a steric governor,

preventing the DnaC ATPase from firing when oligomerized by DnaB to prevent futile

cycling until substrate DNA is bound.

In examining the activity of the Arg-coupler mutant, we also probed the activity of a

DnaC mutant (E176G) encoded by the dnaC810 allele. Purified DnaC810 has been

shown to load DnaB on SSB-coated ssDNA and on D-loops, bypassing the need for the

PriA replication restart protein, and it has been proposed that the DnaB loading activity

of this mutant is mediated by a cryptic DNA- in DnaC (Xu and Marians,

2000). Inspection of the structure shows that Glu176 sits on a loop that connects one-

half of the DNA binding tetrad (Ser177 and Tyr179) to the Walker-B loop of the ATPase

center (Fig. 1C). Interestingly, DnaC810 shows a similar pattern of behavior as the

alanine-substitution of Arg216: DnaB-dependent but ssDNA-independent ATPase

activity, and near wild-type efficiency in loading DnaB onto DNA and activating the

helicase for fork unwinding (Fig. 7). However, unlike the R216A mutant, the dnaC810

allele supports cell viability (Makowska-Grzyska and Kaguni, 2010; Sandler et al.,

1996). A comparison of structures of the DnaBC complex in DNA-free and -bound

states shows that the Walker-B loop on which Glu176 sits switches between two

conformations (Arias-Palomo et al., 2019). Although a structure of DnaC810 bound to

DnaB is currently unavailable, it seems likely that the substitution of the glutamic acid

with glycine alters the flexibility of the Walker-B region, perhaps mimicking a DNA-

bound conformational state to relieve DnaC of the repressive action of Arg216. The

18 bioRxiv preprint doi: https://doi.org/10.1101/2020.10.21.345918; this version posted October 21, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-NC-ND 4.0 International license.

finding that the DnaC810 can act in a manner similar to that of the arginine-coupler

mutant suggests that tight ATPase control by the loader is necessary to properly

coordinate its activity with pathways that resuscitate stalled replication forks.

Understanding precisely how this action might allow DnaC810 and the R216A mutant to

act independently of replication restart factors awaits further study.

Given the widespread nature of AAA+ proteins, we were curious as to whether the

coupling mechanisms we observe for DnaC might be found in other ATPase family

members. Because AAA+ ATPases recognize diverse types of substrates – single and

double stranded nucleic acids, protein chains – it seemed unlikely that there would be

any consensus to their substrate-recognition sites. However, DnaA, a DnaC paralog

that serves to reorganize and melt origins to initiate replication in bacteria, does retain a

highly conserved arginine in the same position as the Arg-coupler. Sequence analyses

show that two proteins used to catalyze replicative helicase loading in eukaryotes, Orc4

and Orc1, also retain such an element (Fig. 8C). Interestingly, these proteins all belong

to the same ‘initiator’ subgroup, or clade, of AAA+ proteins, suggesting that the coupling

arginine seen in DnaC may play a similar role in regulating substrate-dependent ATP

turnover in these proteins as well. In the case of DnaA, the integrity of the equivalent of

DnaC-Arg216 (Arg281 in E. coli DnaA) has been shown to be dispensable for ATP

binding and melting of oriC DNA, but necessary for forming a sufficiently stable complex

that can support DnaB loading by DnaC (Felczak and Kaguni, 2004); by comparison,

mutation of the neighboring Arg-finger of DnaA, R285A, leads to a failure in unwinding

oriC (Kawakami et al., 2005). These findings echo the retention of certain specific

19 bioRxiv preprint doi: https://doi.org/10.1101/2020.10.21.345918; this version posted October 21, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-NC-ND 4.0 International license.

activities but an overall functional defect seen for DnaCR216A. Interestingly, although

there is no known data on the phenotypic effect of mutating the equivalent residue in

human Orc4 (Arg205), mutation of the analogous residue in Orc1 (Arg666, to Trp) has

been linked to Meier-Gorlin syndrome (Guernsey et al., 2011). It has yet to be

determined biochemically whether Arg-coupler mutations in ORC might also lead to an

unlinking of ATPase activity from DNA binding status as is seen for DnaC or to a

different functional defect.

Other proteins in the initiator clade, such as IstB (a transposition helper protein and

another DnaC paralog), do not possess a clear homolog of the Arg-coupler, nor do

several archetypal AAA+ ATPases of other clades (e.g., clamp loader, classic, pre-

sensor I insert, etc.). A review of several representative structures of these proteins

nonetheless does reveal a potential common theme shared between DnaC, IstB, and

clamp loaders. In addition to blocking access to the reactive γ-PO4 of bound nucleotide,

the Arg-coupler of DnaB-bound DnaC also engages the polar Sensor-I amino acid (Fig.

8D), and releases this interaction when DNA is bound. The Sensor-I amino acid is a

conserved feature of AAA+ proteins and is generally important for ATP hydrolysis

(Hattendorf and Lindquist, 2002; Neuwald et al., 1999b; Steel et al., 2000). In the case

of IstB – whose ATPase is primed by DNA binding but not actuated until after it engages

its client IstA transposase – a DNA-bound structure of the protein shows that the Arg-

coupler is replaced with a small amino acid (e.g., alanine), but that a bulky amino acid

(tyrosine) on the Walker-B loop occupies the same general position to occlude the g-

PO4 moiety and remodel the Sensor-I loop (Fig. 8E) (Arias-Palomo and Berger, 2015).

20 bioRxiv preprint doi: https://doi.org/10.1101/2020.10.21.345918; this version posted October 21, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-NC-ND 4.0 International license.

Clamp loaders, whose AAA+ clade is closely-related to that of the initiators, also lack

the upstream Arg-coupler; however, in the presence of a client clamp – which is

necessary but insufficient for stimulating ATPase activity on its own – access to the g-

PO4 is again occluded by a Sensor-I interaction with a positively charged residue on an

adjacent helix (Fig. 8F). A comparison of clamp loader structures reveals that an

arginine or lysine in this highly-conserved central helix can, upon DNA binding, swing

out to release the sensor-I, thereby allowing the Arg-finger to access the g-PO4

(Bowman et al., 2004; Gaubitz et al., 2020; Simonetta et al., 2009). An interesting

distinction of initiators and clamp/helicase/transposase loaders compared to other AAA+

systems is that they are switch-like in their behavior, as opposed to acting in a more

processive manner. Restricting access to g-PO4 of ATP and sequestering the Sensor-I

amino acid may thus serve as a general fail-safe in these systems, as it does in DnaC,

to allow an active ATPase center to be assembled while avoiding premature hydrolytic

events. Whether other more switch-like AAA+ proteins have evolved similar regulatory

mechanisms remains to be determined.

ACKNOWLEDGEMENTS

We would like to thank current and former members of the Berger lab, in particular

Ernesto Arias-Palomo and Frédéric Stanger, for helpful discussions during the analysis

and preparation of data. This effort was supported by the NIGMS (R37-GM71747 to

JMB and R01-GM098885 to JLK).

21 bioRxiv preprint doi: https://doi.org/10.1101/2020.10.21.345918; this version posted October 21, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-NC-ND 4.0 International license.

AUTHOR CONTRIBUTIONS

Conceptual planning – NP, AJF, VLOM, JMB; data collection – NP, AJF, VLOM, SM;

data analysis and writing – all authors.

22 bioRxiv preprint doi: https://doi.org/10.1101/2020.10.21.345918; this version posted October 21, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-NC-ND 4.0 International license.

EXPERIMENTAL METHODS

Protein expression and purification

6xHis-MBP tagged E. coli dnaC was cloned in a pET28b-derived , while E. coli

dnaB was cloned into pET28b without an affinity tag. All constructs were sequence

verified. Wild-type proteins were expressed in strain C41 (Lucigen) by induction at mid-

log phase (OD600~0.4) with 1 mM IPTG at 37° C for 3h (DnaB) or 2.5h (DnaC). Following

induction, cells were harvested by centrifugation and resuspended in lysis buffer

supplemented with protease inhibitors (for DnaB – 20 mM HEPES-KOH (pH 7.5), 500

mM NaCl, 10% glycerol, 10 mM MgCl2, 0.1 mM ATP, 1 mM β-mercaptoethanol; for DnaC

– 50 mM HEPES-KOH (pH 7.5), 1 M KCl, 10% glycerol, 30 mM imidazole, 10% MgCl2,

0.1 mM ATP, 1 mM β-mercaptoethanol; 1 mM PMSF, 1 μg/mL Pepstatin A, and 1 μg/mL

Leupeptin in both). Resuspended cells were flash-frozen and stored at -80˚ Cfor later use.

For purification, cells containing expressed protein were thawed, lysed by sonication, and

clarified by centrifugation. Clarified lysates containing wild-type DnaB were ammonium

sulfate precipitated (30% wt/v), followed by resuspension of the pellet in 100 mM NaCl Q

Buffer (100 mM NaCl, 20 mM HEPES-KOH (pH 7.5), 10 % glycerol, 10 mM MgCl 2, 0.01

mM ATP, 1 mM b-mercaptoethanol, and protease inhibitors) and passed over an HiTrap

Q HP anion exchange column (GE Healthcare). Peak fractions were pooled, concentrated

in Amicon Ultra Centrifugal Filters (30,000 MWCO, Millipore) and further purified by gel

filtration on a HiPrep 16/60 Sephacryl S-300 column (GE Healthcare) in buffer containing

20 mM Tris-HCl (pH 8.5), 800 mM NaCl, 10% glycerol, 5 mM MgCl2, 1 mM β-

mercaptoethanol, 0.1 mM ATP, 1 mM PMSF, 1 μg/mL Pepstatin A, and 1 μg/mL

23 bioRxiv preprint doi: https://doi.org/10.1101/2020.10.21.345918; this version posted October 21, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-NC-ND 4.0 International license.

Leupeptin. Peak fractions were then concentrated and applied to a second sizing column

run in buffer containing 100 mM NaCl rather than 800 mM NaCl.

Purification for WT DnaC from clarified lysates was performed using Ni-Sepharose (GE

Healthcare) affinity resin, followed by TEV protease incubation (to remove the affinity tag)

and a second Ni-Sepharose step. The flow-through from this step was concentrated in an

Amicon Ultra Centrifugal Filters (10,000 MWCO, Milipore) and applied to a HiPrep 16/60

Sephacryl S-200 gel filtration column (GE Healthcare) in buffer containing 50 mM HEPES

(pH 7.5), 500 mM KCl, 10% glycerol, 10 mM MgCl2, 0.1 mM ATP, 1 mM β-

mercaptoethanol, 1 mM PMSF, 1 μg/mL Pepstatin A, and 1 μg/mL Leupeptin.

To overcome expression problems arising from toxicity, mutants of DnaB and DnaC were

expressed into the periplasm. Wildtype dnaB and dnaC genes were cloned as 6xHis-MBP

fusions with an N-terminal periplasmic localization signal, and point mutations were

introduced into these by QuickChange site-directed mutagenesis. All constructs

were sequence verified. Mutant proteins were expressed in strain C41 (Lucigen) by

induction at mid-log phase (OD600 ~0.4) with 0.5 mM IPTG overnight at 16˚ C. Following

induction, cells were harvested by centrifugation, resuspended into the appropriate lysis

buffer for the wild type proteins, flash frozen, and stored at -80˚ C for later use. Purification

protocols followed those of wild-type proteins, except that the DnaBKAEA double mutant

was purified using sequential Ni-Sepharose (GE Healthcare) and amylose (New England

Biolabs) affinity resins, followed by TEV protease incubation (to remove the affinity tag)

and a second Ni-Sepharose step.

24 bioRxiv preprint doi: https://doi.org/10.1101/2020.10.21.345918; this version posted October 21, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-NC-ND 4.0 International license.

One mutant, DnaCE170Q (a Walker-B mutant), did not express using a periplasmic

expression strategy. Expression of this construct was eventually worked out using the

BL21AI expression strain (Invitrogen) grown in MDG-M9ZB medium (Studier, 2005).

Freshly transformed cells were plated on 2xYT plates with 1% glucose to minimize leaky

expression. Starter cultures were grown overnight in MDG minimal media, and

subsequently grown in M9ZB to O.D. ~2. Protein expression was induced with 0.2%

Arabinose overnight at 16°C. After expression, the material was processed as described

for other DnaC mutants.

The purity of all DnaB and DnaC proteins throughout different purification steps was

assessed by SDS-PAGE. The concentration of the purified factors was determined using

ND-1000 UV-Vis Spectrophotometer (Thermo Fisher) by measuring absorbance at 280

nm. Following purification, proteins were aliquoted, flash frozen in liquid nitrogen, and

stored at –80° C. Aliquots were thawed on ice for in vitro assays and used once; any

remaining protein was discarded.

ATPase activity assay

Measurement of DnaC ATPase activity was carried out as described previously (Arias-

Palomo et al., 2019). Briefly, coupled ATP-hydrolysis assays were performed in 40 mM

HEPES (pH 7.5), 10 mM MgCl2, 5 mM DTT, and 0.1 mg/ml BSA. A single Walker-A

mutant of (K236A) proved to have high residual ATPase activity background (not shown),

so a double mutant was made in conjunction with catalytic glutamate substitution (E262A,

25 bioRxiv preprint doi: https://doi.org/10.1101/2020.10.21.345918; this version posted October 21, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-NC-ND 4.0 International license.

referred to as ‘DnaBKAEA’). To assess DnaC activity, 5 µM of purified protein was mixed

with 5.25 µM DnaBKAEA and 2.1 µg M13 ssDNA, 0.5 mM NADH, and 15 mM

phosphoenolpyruvate in presence of a 4% v/v pyruvate /lactate dehydrogenase

enzyme mix (Sigma). Reactions were initiated by the addition of ATP to a final

concentration of 25 µM - 6.4 mM, and the absorbance of the reaction was monitored at

340 nm in a 96-well, half-area plate (Corning) using a CLARIOstar (BMG Labtech)

microplate reader at 37° C. An NADH standard curve was used to convert A340 to NADH

concentration, which in turn was used to calculate the rate of NADH loss that linearly

correlates to ATP-hydrolysis; initial rates were then calculated and plotted against ATP-

concentration. The data were fit to Michaelis-Menten equation using Prism to estimate

kcat and Km values. The experimental data presented are from three independent

experiments.

DnaB loading assay

We utilized a gel-based assay with M13 ssDNA as a substrate to assess the efficiency of

DnaB loading (Fang et al., 1999). Details of the assay have been described previously

(Arias-Palomo et al., 2019). Briefly, purified DnaBWT or DnaBKAEA was mixed with

equimolar amounts of wildtype or mutant DnaC to a final volume of 100 µL and a final

concentration of 10 µM in presence of 40 mM HEPES (pH 7.5), 10 mM MgCl2, 5 mM DTT,

0.1 mg/ml BSA, 5 mM ATP, and 1 µg M13 DNA. Reactions were incubated for 15 min at

30 °C and then applied to a Bio-Gel A-1.5m (Bio-Rad) resin packed in a 0.7X15cm Flex-

Column (Kimble). 250 µl fractions were collected by hand at 20˚ C, mixed with SDS-PAGE

gel-loading dye, and resolved on 12% Tris-Glycine SDS-PAGE gels. Gels were silver-

26 bioRxiv preprint doi: https://doi.org/10.1101/2020.10.21.345918; this version posted October 21, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-NC-ND 4.0 International license.

stained, imaged on an Gel Doc EZ gel-documentation system (BioRad), and the amount

of DnaB in each fraction estimated using Image Lab (BioRad). Helicase loading efficiency

was calculated as the ratio of DnaB that eluted with ssDNA in the void volume of the

column compared to the total amount of DnaB present in all fractions (Fig S1B). A few

DnaC mutants (e.g., N203A or R220A) gave rise to a population of ssDNA-associated

DnaB that migrated anomalously through the gel filtration matrix with a markedly delayed

elution profile. These fractions were considered to contain aberrant or misloaded

complexes and were not included in calculating the amount of DnaB loaded compared to

wild-type DnaC (which never displayed such behavior) – the DnaB present in these

fractions was used for calculating the total amount DnaB only. The experimental data

shown reflect the average of three independent experiments. Graphs were generated in

Prism (GraphPad).

DNA Fork Unwinding Assay

Fork unwinding assay was performed as described previously (Arias-Palomo et al., 2013),

with minor modifications. Briefly, HPLC-purified DNA oligonucleotides were purchased

from Integrated DNA Technologies. The forked DNA substrate was formed by annealing

20 μM 5¢ -Cy3-labeled oligonucleotide (5¢-Cy3-TACGTAACGAGCCTGC(dT)25-3¢) with a

1.2 molar excess of a 3¢-black quencher (BHQ)-labeled strand (5¢-(dT)25-

GCAGGCTCGTTACGTA-BHQ2-3¢) in annealing buffer (20 mM Tris-HCl (pH 7.5), 10 mM

MgCl2, and 20 mM KCl). A control “captured” fork substrate with 100% maximum

fluorescence signal was formed similarly to the forked DNA substrate, but that substituted

the BHQ strand with an unlabeled capture strand (complementary to the base-paired

27 bioRxiv preprint doi: https://doi.org/10.1101/2020.10.21.345918; this version posted October 21, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-NC-ND 4.0 International license.

region of the Cy3-labeled oligo, 5¢-GCAGGCTCGTTACGTA-3¢). The captured-fork

control was used to normalize signal between experiments. For the reactions, helicase

and loader were premixed on ice, followed by the addition of substrate DNA. This reaction

mixture was pre-warmed to 37 °C; in parallel, the single-stranded capture oligo DNA was

mixed with ATP, and pre-warmed to 37°C separately. The ATP/capture-strand mixture

was then added to the pre-warmed reaction mixture, pipetted gently to mix, and then three

20 μL aliquots were immediately pipetted into and read out from a 384-well black plate

using a CLARIOstar (BMG Labtech) microplate reader at 37 °C. The final reactions

containing 200 nM DnaB hexamer (with or without 1.2 μM loader monomer), 100 nM fork

substrate, 200 nM capture strand, and 1 mM ATP were monitored for fluorescence

increase at 37 °C for 26 min. The final assay buffer consisted of 20 mM HEPES-KOH (pH

7.5), 5 mM acetate, 50 mM potassium glutamate, 5% glycerol, and 0.2 mg/mL

BSA. Reads from triplicate wells were averaged and analyzed in Excel (Microsoft) and

normalized graphs were generated in Prism (GraphPad). Presented data are from three

independent repeats.

DnaC-ssDNA binding assays

Fluorescence anisotropy was used to quantitate DNA-binding efficiency of DnaC variants.

DnaC interaction assays with ssDNA were performed as previously described (Arias-

Palomo et al., 2019; Mott et al., 2008). 0-50 μM DnaC was first mixed in a high salt buffer

(50 mM HEPES (pH 7.5), 500 mM KCl, 10% (v/v) glycerol, 10 mM MgCl2) before being

rapidly diluted four-fold into binding buffer containing 50 mM HEPES (pH 7.5), 10% (v/v)

glycerol, 5 mM MgCl2, 2 mM ATP, 2 mM DTT, 0.1 mg/ml BSA, and 10 nM fluorescein-

28 bioRxiv preprint doi: https://doi.org/10.1101/2020.10.21.345918; this version posted October 21, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-NC-ND 4.0 International license.

labeled dT25 oligonucleotide (IDT). Protein from the high salt buffer was diluted in the

binding reaction to bring the final salt concentration to 125 mM KCl. Aliquots of the

samples were transferred to prewarmed 384-well, black small volume polystyrene plates

(Greiner Bio-One). The plates were incubated for 10 min and then read in a CLARIOstar

(BMG Labtech) microplate reader at 37° C. Averaged delta-anisotropy values were

plotted against increasing DnaC concentrations. Data were fit to Hill equation for specific

binding using Prism. The experimental data shown are from three independent repeats.

PriC-dependent unwinding assays

Reactions were performed in triplicate using DNA substrates as previously described

(Wessel et al., 2013) with the final monomeric concentrations of components as follows:

67 nM SSB, 120 nM DnaB, 400 nM DnaC (or variant), and 40 nM PriC. Briefly, a forked

DNA substrate (60 bp duplex, 38 nt 3' and 5' overhangs) was made by annealing two

oligonucleotides, one of which is labeled with 32P. The substrate was preincubated with

SSB (when included) for 3 min at 25 °C. Reactions were initiated by the addition of DnaB,

DnaC (or variant), and PriC (when included) and incubated at 37 °C for 30 min. Unwinding

was terminated by the addition of 20 mM EDTA, 0.5% SDS, 0.2 mg/ml of proteinase K,

and 2.5 ng/ul of oligonucleotide 3L-98 (final concentrations) and incubated at 37 °C for

30 min. Samples were resolved on a 10% native polyacrylamide gel. The gel was then

fixed in 10% methanol, 7% acetic acid, and 5% glycerol, dried, and exposed to a

phosphorimager screen and imaged on a Typhoon FLA 9000. Band intensities were

quantified using ImageQuant (GE Healthcare) and percent unwinding was determined by

dividing the intensity of the single strand product band by the total intensity in the lane.

29 bioRxiv preprint doi: https://doi.org/10.1101/2020.10.21.345918; this version posted October 21, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-NC-ND 4.0 International license.

FIGURE LEGENDS

Fig. 1: DnaC functional elements. A) Domain organization of DnaC. B) Structure of an

ssDNA-bound DnaBC complex from E. coli shown as a cartoon view (Arias-Palomo et

al., 2019). DnaB is colored blue (NTD) and purple (RecA ATPase domain), DnaC

subunits are alternatingly orange and yellow, and ssDNA is red. C) Close up cartoon

view of the DnaC active site showing elements mutated in this study. One protomer is

colored orange, the other yellow; DNA is red. Motifs: WA – Walker A (Lys112); WB –

Walker B (E170); SI – Sensor I (Asn203); RC – Arginine Coupler (Arg216); RF –

Arginine Finger (Arg220); SII – Sensor II; 810 – Glu176; Tetrad – Lys143, Phe146,

Ser177, Tyr179.

Fig. 2: Effects of AAA+ motif mutations on DnaC ATPase and DnaB loading activities.

A) Comparison of ATPase activity between wild-type DnaC and Walker-A, Walker-B,

Sensor-I, Arg-finger, and Sensor-II mutants. B) Silver-stained SDS-PAGE analysis of

fractions from a representative DnaB-loading reaction containing DnaC and M13 ssDNA

that has been passed over a sizing column. Lanes where DnaB co-migrates with DNA

indicate loading. Some DnaC remains bound to loaded DnaB under these conditions. C)

Comparison of DnaB loading efficiencies for different DnaC ATPase mutants. HBD –

helicase binding domain of DnaC. Error bars on this an all other bar graphs reflect the

standard deviation in loading from at three independent experimental replicates.

Asterisks on this and all other bar graphs correspond to P-values from one-way ANOVA

tests of all mutants against WT DnaC: ‘****’ – P<0.0001; ‘***’ – P<0.001; ‘**’ – P<0.01; ‘*’

– P< 0.05; ‘ns’ – not significant.

30 bioRxiv preprint doi: https://doi.org/10.1101/2020.10.21.345918; this version posted October 21, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-NC-ND 4.0 International license.

Fig. 3. DnaB loading is impacted by helicase ATPase status, and can be reversed by

DnaC. A) DnaB loading is compromised in an ATPase-deficient mutant of DnaB. B) DnaB

loaded on M13 ssDNA is large salt resistant, consistent with structural studies showing

that helicase ring topologically closes around DNA in the presence of DnaC. C) DnaC can

actively unload DNA in an ATP-dependent manner. The inclusion of ATP in the column

running buffer reverses stable DnaB loading, whereas ADP does not.

Fig. 4. Effect of DnaC ATPase status and 3’-end accessibility on the stimulation of DNA

unwinding by DnaB. A) The addition of a 3’-end block to a DNA fork partially stimulates

DnaB-dependent fork unwinding compared to when DnaC is present. The percent of DNA

strand separation as measured by a gain in fluorescence intensity is plotted as a function

of time. B) The isolated DnaC HBD readily stimulates unwinding of an unblocked fork

compared to wildtype DnaC. C) The ATPase activity of DnaC is necessary for stimulating

fork unwinding by DnaB. Error bars on this an all other unwinding curves reflect the

standard deviation from different loads of the same reaction in a single experiment.

Fig. 5. Effect of DNA binding on DnaC function. A) WebLogos (Schneider and Stephens,

1990) showing the conservation of the Lys143/Phe146 and Ser177/Tyr179 regions of

DnaC. B) Ribbon and stick representation showing how ssDNA binds to DnaC (Arias-

Palomo et al., 2019). C-E) Bar graphs showing that Lys143 of DnaC is essential to: C)

DnaC ATPase function, D) DnaB loading, E) stimulation of DNA fork unwinding by DnaB.

31 bioRxiv preprint doi: https://doi.org/10.1101/2020.10.21.345918; this version posted October 21, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-NC-ND 4.0 International license.

Fig. 6. Functional effect of a second conserved arginine adjacent to the Arg-finger on

DnaC. A) WebLogo (Schneider and Stephens, 1990) showing the conservation of

Arg216, which sits four amino acids away from the DnaC Arg-finger (Arg220). B-D) Bar

graphs showing the effect of R216A and R216E mutants on: B) ssDNA- and DnaB-

dependent stimulation of DnaC ATPase activity, C) DnaB loading by DnaC, and D) DnaC-

stimulation of DNA fork unwinding by DnaB. E) Bar graph showing that the R216A

mutation can override the loss of DnaC ATPase activity imparted by the disruption of

Lys143 when DnaB is present.

Fig. 7. Loss of the ssDNA dependency on DnaC ATPase activity correlates with an ability

to bypass a need for the replication restart factor, PriC, in unwinding SSB-coated DNA.

A-C) The DnaC810 replication restart bypass mutation phenocopies a loss of the Arg-

coupler. Bar graphs show the effect of the DnaC810 E176G mutation on: A) DnaB

loading, B) DnaB-dependent DNA fork unwinding, and C) ssDNA- and DnaB-stimulated

ATPase activity of the loader. D) The Arg-coupler mutant of DnaC promotes the DnaB-

mediated unwinding of an SSB-coated fork in a PriC-independent manner. Error bars

reflect the standard deviation from three different experiments.

Fig. 8. Molecular mechanism of the DnaC Arg-coupler and the presence of potential

functional analogs in related switch-type AAA+ ATPases. A,B) Schematics and structures

showing how the conformational status of the Arg-coupler switches between apo and

DNA-bound states to respectively occlude or allow access to the g-PO4 moiety of

nucleotide in the DnaC ATPase site. C) WebLogo (Schneider and Stephens, 1990)

32 bioRxiv preprint doi: https://doi.org/10.1101/2020.10.21.345918; this version posted October 21, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-NC-ND 4.0 International license.

showing an analog of Arg216 (red dotted box) is present next to the Arg-finger (cyan

dotted box) of DnaA, Orc1, and Orc4., which sits four amino acids away from the DnaC

Arg-finger. D-F) The Sensor I residue of IstB and (RFC) is engaged

by a conserved side chain in unstimulated ATPase states. DnaC PDB 6QEL; IstB PDB

5BQ5; RFC PDB 1SXJ.

Fig. S1. Silver-stained SDS-PAGE analysis of sizing-column fractions from DnaB loading

reactions containing: A) no DnaC, B) the DnaC HBD, C) a DnaC SI mutant, D) a DnaC

WB mutant, E) a DnaC RF mutant, F), a DnaC SII mutant, and G) a DnaC WA mutant.

Red arrowheads indicate early fractions where DnaB co-migrates with ssDNA, indicating

loading.

Fig. S2. Silver-stained SDS-PAGE analysis of representative sizing-column fractions

from DnaB loading reactions containing: A) WT DnaC and an ATPase-deficient (KAEA)

mutant of DnaB, B) WT DnaB and DnaC, but where 250 mM NaCl was used in the

column running buffer, C) WT DnaB and DnaC, but where ATP was added to the

column running buffer, D) WT DnaB and DnaC, but where ADP was added to the

column running buffer, and E) WT DnaC and the KAEA ATPase-deficient mutant of

DnaB, but where ATP was added to the column running buffer. All gels except that

shown in (B) reflect experiments where columns were run in 100 mM NaCl. Red

arrowheads mark where DnaB co-migrates with DNA, indicating loading.

33 bioRxiv preprint doi: https://doi.org/10.1101/2020.10.21.345918; this version posted October 21, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-NC-ND 4.0 International license.

Fig S3. Effect of DnaC ATPase status on the stimulation of DnaB unwinding activity.

Graphs show the gain in fluorescence intensity of a specified DnaC variant as a

compared to the ability of DnaB to unwind a model fork substrate in absence of

presence. DnaC tested are: A) Walker-A, B) Walker-B, C) Sensor I, D) Arg-finger, and

E) Sensor II.

Fig. S4. DnaC residues seen to engage ssDNA structurally are important for ssDNA

binding, helicase loading, and stimulating DNA unwinding by DnaB. A) Unwinding

activity for F146., B) Unwinding activity for F146A Y179A. C) Defect in DnaCK143E DNA-

binding function as measured by its ability to bind a fluorescein-labeled dT25

oligonucleotide. D) Unwinding activity for K143E DnaC. E) Silver-stained SDS-PAGE

analysis of sizing-column fractions from a representative DnaB loading reaction with

DnaCK143E.

Fig. S5. Effect of coupler amino acids on DnaC function. A) Representative unwinding

assay comparing the stimulatory activity of two different Arg-coupler mutants (R216A

and R216D) to wild-type DnaC. B,C) Representative loading SDS-PAGE gel assays

showing the ability of two different Arg-coupler mutants (R216A and R216D) to deposit

DnaB on M13 ssDNA. D, E) Representative unwinding assay comparing the stimulatory

activity of a double Arg-coupler/ssDNA-binding mutant (R216A/K143E, panel D) or the

DnaC810 mutant (Panel E) to wild-type DnaC. F) Representative silver-stained SDS-

PAGE analysis of sizing-column fractions from DnaB loading reactions containing

DnaC810. G) Representative, gel-based unwinding assay showing that the DnaC810

34 bioRxiv preprint doi: https://doi.org/10.1101/2020.10.21.345918; this version posted October 21, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-NC-ND 4.0 International license.

and Arg-coupler mutants effectively promote the unwinding of an SSB-coated DNA fork

substrate in the absence of PriC.

Movie S1. Morph movie going from Apo (DNA-free) to DNA-bound states of DnaB-

bound DnaC, showing a close-up of the DnaC ATPase site and the movement of the

Arg-coupler. Conserved catalytic amino acids colored as per Fig. 1C.

35 bioRxiv preprint doi: https://doi.org/10.1101/2020.10.21.345918; this version posted October 21, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-NC-ND 4.0 International license.

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53